Integration of native and engineered photosynthetic microorganisms into artificial assemblies for the development of promising biosensors targeted to environmental monitoring

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DI~ARnMENTODELL' AGRICOLTURA, fOREStE, NATuRA ED ENERGIA

COiSO DI DonORAlO DI' RICERCA

_BIOIECNOLOGIE VEGETALi,_~_ - XXIV Ciclo.

INTEGRATlo'N OF NATIVE ANO ENGINEERED PHOTOSYNTI:IETIC MICROORGANISMSJNJO A,RTIFICIALASSEMBLIES FOR THE DEVELOPMENT OF PROMISING BIOSENSORS TARGETED TO

ENVIRONMENTAL MONITOIUNG

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OO.ordinafore:Prof. SfeJpnia Mosci

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Tutor: Dott.ssa' Maria Teresa Giarçli

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1 TABLE OF CONTENTS 1.Introduction ... 4 1.1 Biosensor description ... 4 1.1.1 Transducer System ... 7 1.1.1.1 Electrochemical Biosensors ... 7 1.1.1.1.1 Conductimetric biosensors ... 7 1.1.1.1.2 Potenziometric biosensors ... 8 1.1.1.1.3 Amperometric biosensor ... 9 1.1.1.1.3.1 Cyclic Voltammetry ... 11

1.1.1.2 Other transducer system ... 11

1.1.1.2.1 Piezo-electric biosensor ... 11

1.1.1.2.2 Optical biosensors ... 12

1.1.2 Immobilization methods in electrochemical biosensors ... 13

1.1.3 Biosensors to detect pesticides ... 13

1.2 Photosynthesis ... 15

1.2.1 Higher plants and green algae photosynthesis ... 15

1.2.2 Purple Bacteria ... 19

1.3 Photosynthesis biosensor ... 23

1.4 Aim and develop of PhD project... 25

1.4.1 Development of biosensor whole cell based C. reinhardtii ... 25

1.4.2 Development of Chimera RC Protein ... 27

2.Experimental Section ... 28

2.1 Development of biosensor based on whole cells of Chlamydomonas reinhardtii ... 28

2.1.1 Chlamydomonas reinhardtii growth ... 28

2.1.2 Chlamydomonas reinhardtii characterization ... 29

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2.1.2.2 Cell density. ... 29

2.1 2.3 Fluorescence yield. ... 29

2.1.2 Algae cells Nafion® immobilization on screen-printed electrodes ... 30

2.1.3 Algae cells Alginate immobilization on screen-printed electrodes ... 30

2.1.4 Biosensor instrument setup ... 31

2.1.5 Amperometric Measure ... 32

2.1.5.1 Electrode Choosing ... 32

2.1.5.2 Setting of Electrochemical Measures (Nafion® immobilization) ... 33

2.1.5.2 Setting of Electrochemical Measures (Alginate immobilization) ... 34

2.1.5.2.1 Cyclic voltammetry tests on screen-printed biosensors. ... 34

2.1.5.2.2 Chronoamperometry tests on screen-printed biosensors ... 34

2.1.5.2.3 Chronoamperometry tests: storage stability and stability during measurements. ... 34

2.1.5.2.4 Chronoamperometry tests: evaluation of herbicide inhibition ... 35

2.1.6 Alternative measurements for the evaluation of herbicide inhibition ... 35

2.1.6.1 Oxygen Evolution Rate tests ... 35

2.1.6.2 Fluorescence tests. ... 36

2.2 Chimera Creation ... 36

2.2.1 Bioinformatics analysis and future RC genetic manipulation ... 36

2.2.2 Genetic engineering on R. sphaeroides and C. reinhardtii organisms ... 37

2.2.3 Photosynthetic membranes isolation ... 41

2.2.4 Absorbtion spectroscopy characterization... 42

2.3 Protocols Section... 43

2.3.1 Protocol 1 TAP Production... 43

2.3.2 Protocol 2 Growth Rhodobacter sphaeroides ... 44

2.3.2.1 Appendix A : M22+ medium 10x concentrate ... 45

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2.3.2.3 Appendix C: M22+ liquid medium ... 47

2.3.2.4 Appendix D: 104 Vitamins ... 47

2.3.2.5 Appendix E: M22+ agar ... 48

2.3.2.6 Appendix F: Antibiotics ... 48

2.3.2.7 Appendix G: Glycerol Stocks ... 49

2.3.3 Protocol 3: Wizard® Plus Minipreps DNA Purification System -Promega ... 49

2.3.4 Protocol 4: Wizard® SV Gel and PCR Clean-Up System ... 50

2.3.5 Protocol 5 TAE... 51

2.3.6 Protocol 6 TBE... 52

2.3.7 Protocol 7 Competent Cells ... 52

2.3.8 Protocol 8 Luria Bertani (LB) Medium ... 53

Protocol 9 Mating Protocol ... 53

2.3.10 Protocol 10 Membranes – sucrose cushion ... 54

3.1 Development of biosensor ... 56

3.1.1 Nafion immobilization methods ... 57

3.1.1.2 Viability of immobilized algae in chronoamperometry: "half-life” measurements. .. 58

3.1.2 Alginate immobilization method ... 60

3.1.2.1 Cyclic voltammetry: investigation on the electrochemical reaction at electrode surface ... 61

3.1.2.2 Chronoamperometry: the principles of the herbicide detection. ... 63

3.1.2.3 Biosensor performances in the absence of herbicides. ... 65

3.1.2.4 Biosensor performances in the presence of herbicides. Suggestions about QC site . 67 3.1.2.5Comparison with analogous biosensing devices and conventional methods. ... 71

3.1.2.6Preliminary results with C.reinhardtii mutant ... 72

3.2 Engineering of Rhodobacter sphaeroides Reaction Center ... 73

3.2.1 Chimera Creation to biosensoristic purpose ... 73

4 3.2.3 Chimera creation ... 78 4.Conclusion... 81 5.References ... 84

1.Introduction

Biosensors are devices which use a biological recognition element retained in direct spatial contact with transduction system (IUPAC definition) [1]. Biosensors can be straightforwardly depicted as devices that convert a physical or biological event into a measurable signal [2] they are composed of a biosensing element (i.e. enzyme, tissue, living cell) that provides selectivity and a transducer that converts chemical signal into a processable signal [3]. Specifically, biosensor consists of three parts: the first element is the biomediator (a biomimic or biologically derived material e.g. tissue, microorganisms, organelles, cell receptors, enzymes, antibodies, nucleic acids, and biological sensitive elements created with genetic engineering), the second element is the transducer (physicochemical, optical, piezoelectric, electrochemical, etc.) that transforms the signal resulting from the analyte‘s interaction with the biological element into a signal that can be measured and quantified; the third element is the associated electronics or signal processor, responsible of the results visualization in a user-friendly way [4]. Biosensors require immobilization of biomediator to the surface of the sensor (metal, polymer or glass and other materials) using physical or chemical techniques (Figure 1)

5 Figure 1.1 Biosensor scheme

Looking at the past it is clear that the concept of biosensor has evolved. The first example of biosensor was illustrated by entrapping the enzyme glucose oxidase in a dialysis membrane over an oxygen probe. Later, the same term "enzyme electrode" was used to describe a similar device where again the enzyme glucose oxidase was immobilized in a polyacrylamide gel onto a surface of an oxygen electrode for the rapid and quantitative determination of glucose. Since the beginning of the ‗80s, several authors started to prove the concept of biosensor as a system in which an enzyme is coupled to electrochemical-optical sensors. In the electrochemical community at that period the research on Ion Selective Electrodes (ISE) was very active with the idea to extend the range of sensors, and even to non ionic compounds, like glucose. Enzymes, multiple enzymes, organelles, bacteria, specialized biological tissues containing specific enzymes were coupled to potentiometric or amperometric, then optical, thermometric, piezoelectric devices, etc. Recently, the concept evolved again in the attempt to replace or mimic the biological material with synthetic chemical compounds, such as MIPs (Molecular Imprinted Polymers). Enzymes and all biological elements based on the enzymes represent the classes of what is now called "catalytic elements". The other important class is represented by the "affinity elements", namely antibodies, lectins, nucleic acids (DNA and RNA) and recently also synthetic ligands. The exploitation of the selectivity of the biological element is the "driving force" of the biosensor [5].

Catalytic events or affinity events have not the same scheme of transduction. If the biological recognition element present in the sensing layer is an enzyme or generally a biocatalyst, a reaction takes place in the presence of the specific target analyte and an increasing amount of co-reactant or product is consumed or formed, respectively, in a short time depending on the turnover. In this scheme, the amplification step is inherent and a large chemical amount can be obtained from the sensing layer. In contrast, the use of antibodies for the detection of antigens does not involve an amplification stage and the "affinity" reaction should be amplified in order to have a clearly readable transduction. We have two possibilities, one is the use of a bioconjugate involving a bound enzyme, like in the classical ELISA test; the second is the inherent amplification given by the mass of the biological element involved where a piezoelectric device (sensitive to mass) can detect minute amount of large proteins (like antibodies) if they are attracted on the surface of the sensor. With the same scheme, surface plasmon resonance can be sensitive to a small amount of large molecule reacting at the surface of the electrode. The main problem of the biological system, catalytic or affinity, is the associated fragility and the operational activity. Most proteins have an

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optimal pH range in which their activity is maximal; this pH range should be compatible with the transducer. Moreover, most of the biological systems have a very narrow range of operating temperature (15÷40°C). The most important problem and main drawback for industrial exploitation is the short lifetime associated with the biological elements. Lifetime of at least months or few years is the prerequisite for a suitable market and the fragility of the assembled systems has always limited the diffusion of biosensors into the market.

In practice the number of biosensors that are likely to be produced and used as ―real‖ commercial devices is very low and the commercial applicability has to be deeply evaluated. Several companies put on the market pre-activated membranes suitable for the immediate preparation of any bioactive membrane and this appeared as a real improvement at least for the easy use of enzyme sensors. The removal of interference has been also another important aspect for the wide use of biosensors in industrial processes. These two main problems have been solved by using multilayer membranes, such as those developed by Yellow Springs Instrument Co. (for glucose or lactate electrodes), where the enzyme is sandwiched between a special cellulose acetate membrane and a polycarbonate nucleopore membrane. The main role of the membrane is to prevent proteins and other macromolecules from passing into the bioactive layer. Cellulose acetate membrane allows only molecules of the size of hydrogen peroxide to cross and contact the platinum anode, thus preventing interference from ascorbic acid or uric acid, for example, at the fixed potential. Such configuration has been used by several researchers in the biosensor assembly. But at the same time several recipes of immobilization of enzymes were published and several laboratories developed their own procedures for immobilizing the biological element, sometimes also patented. One approach was also the development of disposable sensor, based on combination of screen printed electrochemical sensors with enzyme adsorbed on the electrode surface. The use of the sensor just for one measurement limited the use of complicated immobilization procedures to as simple as possible, like the physical adsorption on carbon surface. This electrode surface acted as a sponge, and the large protein was easily immobilized even if the bond was weak. This approach is useful only for a quick and rapid measurement.

Concerning immobilization technology, many other methods have been tested to improve bio-receptor stability and to conserve its bioactivity. The most widely used methods of immobilization are: (i) physical or chemical adsorption on a solid surface; (ii) covalent binding; (iii) entrapment within a membrane, surfactant matrix, polymer or microcapsule; (iv) cross-linking between molecules [6].

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The transducer varies depending on the measured parameter; electrochemical, optical, mass and thermal signals are the most common [7]. The measured parameters may be converted into an electrical signal, depending on the type and application of the biosensor. The biosensor advantages are simplicity, small size, robustness, low cost, ability to generate reliable and continuous information, high selectivity and sensitivity, depending on the biomediator type and immobilization methods. This innovative technology represents a valid support to other efficient analytical methods (HPLC, or GC-MS etc.), and finds for instance useful applications in pre-screening analyses.

Biosensors have various application fields: 1) medical applications e.g. glucose monitoring in diabetes patients [8] and other medical health related targets [9] ; 2) environmental applications, e.g. detection of pesticides and river water contaminants [10]; 3) remote sensing of airborne bacteria, e.g. in counter-bioterrorist activities [11]; 4) detection of pathogens [12] 5)determination of toxic levels of substances before and after bioremediation, and of drug residues in food, such as antibiotics and growth promoters, particularly meat and honey [13], 6) analysis of food quality (e.g. phenols in wine, tea and oil) [14].

1.1.1 Transducer System

The current concept of biosensor date back to 1962 thanks to Leland C. Clarke Jr and his collaborators work [15] proposing an enzymes us biomediator immobilized on a sensor using an electrochemical measurement. The electrochemical biosensors could be divided into three main classes conductimetric, potenziometric, amperometric ones.

1.1.1.1 Electrochemical Biosensors

1.1.1.1.1 Conductimetric biosensors

The conductimetric principle of measurement is widely applicable to chemical systems, because many chemical reactions produce or consume ionic species and thereby alter the overall electrical conductivity of the solution, its resistance being determined by migration of all ions that are present. The principle of operation of a conductimetric biosensor involves the application of an electric field across a pair of microelectrodes surrounded by an electrolyte or a buffered solution containing an enzyme immobilized on a sensor and the species to be detected. An electric field is generated by a sinusoidal voltage waveform across the electrode to minimize or to eliminate the undesirable electric process and to overcome the problems related to temperature variation in the single cell device. An example of conductimetric biosensors is the ―urea biosensor‖, where the urease enzyme

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is immobilized on a chip to detect urea, it catalyzes the decomposition of urea which produce bicarbonate ions (as show below in Reaction R1.1) increasing the conductivity of tested solution [16].

R 1.1) NH2CONH23H2OUrease  2NH 4HCO3OH

1.1.1.1.2 Potenziometric biosensors

Potentiometric biosensors use an Ion-selective electrode (ISE) to transduce a biological reaction into an electrical signal, the reactions involving the release or absorption of ions that may be utilised by potentiometric biosensors. An immobilized enzyme layer catalyses the biological reaction which generates or absorbs ions, when the reaction involves H+ ions, a pH meter is the ISE used.

The potential that develops across an ion-selective membrane separating two solutions is measured at virtually zero current, meaning high impedance and no interference to the reaction [17].

A biorecognition element is immobilized on the outer surface or captured inside the membrane. In the past the pH glass electrode was used as a physicochemical transducer [18]. The Nernst potential of the pH glass electrode is described by the Nicolsky-Eisenman equation, of which the generalized form for ISE is as follows [19] (Equation E 1.1) :

E 1.1)                n 1 i i i a, a a K a ZaF RT E E 0 ln Za/Zi

(E potential, R the universal gas constant, T temperature, F Faraday constant, za followed and zi interfering ion valence, aa activity of measured and ai activity of interfering ion and Ka,i is the selectivity coefficient).

Potentiometric measurements involve, exactly non-faradaic electrode processes, with no net current flow, and operate on the principle of an accumulation of charge density at an electrode surface, resulting in the development of a significant potential at that electrode. This potential is proportional to the logarithm of the analyte activity present in the sample and is measured relative to an inert reference electrode, also in contact with the sample. These sensors are mainly based on field effect transistors (FETs) and were first proposed by Peter Bergveld [20] in 1970 as the ion sensitive field-effect transistor (ISFET). Enzyme-sensitive field-effect transistors (ENFETs) can be fabricated from ISFETs by applying a thin overlayer of enzyme- loaded gel on the ion-selective membrane [21]. Three types of potenziometric biosensors are the morest used: glass electrodes for

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cations (which normally is a pH electrode) pH electrode with a gas permeable membrane, solid state electrodes.

Different example of potenziometric biosensors are developed, for examples o detect urea, penicillin, glucose [22]. as show below in Reaction R 1.2, R 1.3, R 1.4

Pennicillin

R 1.2) penicillin penicilloateH

Glucose

R 1.3) glucoseO2 GOD gluconicacidH2O2

(Gluconic acid change pH, the reaction can be monitored by pH measurement) Urea

R 1.4) CO(NH2)22H2Ourease  2NH2 CO32

(Urea the first analyte detected by biosensor (Guillbaut and Kuan, 1987. Urea is hydrolysed by

ureas)

1.1.1.1.3 Amperometric biosensor

The signal of these biosensors is generated by the electron exchange between the biological system in the bioreceptor layer and one electrode. Generally speaking, when using amperometric biosensors, the analyte undergoes or is involved in a redox reaction that can be followed by measuring the current in an electrochemical cell.

The analytes, or the species involved with it via a (bio)chemical reaction, change the oxidation state at one electrode then the electron flux is monitored and is proportional to the amount of the species electrochemically transformed at the electrode [23].

The most common biosensors are based on chronoamperometric experiments, where the current is monitored as a function of time. Usually a single potential step is used for a single experiment. The analysis of chronoamperometry data is based on the Cottrell equation, which defines the current-time dependence for linear diffusion control (Equation E 1.2):

E 1.2) i=nFACD1/2π -1/2t -1/2

where:

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F= Faraday‘s constant (96,500 C mol-1)

A= electrode area (cm2)

C= concentration (mol cm3)

D= diffusion coefficient (cm2s-1)

That indicates that, under this conditions there is a linear relationship between the current and the 1/square root of time. A plot of i vs. t -1/2 is often referred to as the Cottrell plot [24].

Amperometric biosensors can work in two- or three-electrode configurations. The former case consists of reference and working (containing immobilized biorecognition component) electrodes. The main disadvantage of the two-electrode configuration is a limited control of the potential on the working electrode surface with higher currents, and because of this, the linear range could be shortened. To solve this problem, a third auxiliary electrode is employed. Now voltage is applied between the reference and the working electrodes, and current flows between the working and the auxiliary electrodes. An example of three electrode sensors is Figure 1.2 [25]:

Figure 1.2 Principal Features of Screen Printed Electrode (SPE)

The amperometric biosensors are among the most common biosensors studied both in the past and today, using different types of biological material (enzymes, cells, tissues, proteins, antibodies, nucleic acids) and analytes for clinical use, to detect infection-marker antibodies, acetylcholine, biological oxygen demand, as well as other electrode-attached mediators or to determine simple molecule analytes, such as glutathione, L-alanine and pyruvate, lactate, and cholesterol [26], or for

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food analysis such as lactate, citrate, glutamate, ethanol, ecc. [27], and for environmental analysis to detect toxic substance us pesticide (in the water samples), different type of phenols, surfactants, antibiotics, toxins etc. [28][29].

1.1.1.1.3.1 Cyclic Voltammetry

Electrolysis, cyclic voltammetry (CV) , amperometry and several other techniques might be described as ―active‖ electrochemical methods because the experimenter drives an electrochemical reaction by incorporating the chemistry into a circuit and then controlling the reaction by circuit parameters such as voltage.

Cyclic voltammetry is one of the techniques which electrochemists employ to investigate electrolysis mechanisms, series of voltages are applied to the electrode and the corresponding current that flows monitored, to identify the specific potential of electrochemical reaction.

In typical cyclic voltammetry, a solution component is electrolyzed (oxidized or reduced) by placing the solution in contact with an electrode surface, and then making that surface sufficiently positive or negative in voltage to force electron transfer. In simple cases, the surface is started at a particular voltage with respect to a reference half-cell such as calomel or Ag/AgCl, the electrode voltage is changed to a higher or lower voltage at a linear rate, and finally, the voltage is changed back to the original value at the same linear rate. When the surface becomes sufficiently negative or positive, a solution species may gain electrons from the surface or transfer electrons to the surface. This results in a measurable current in the electrode circuitry. However, if the solution is not mixed, the concentration of transferring species near the surface drops, and the electrolysis current then falls off. When the voltage cycle is reversed, it is often the case that electron transfer between electrode and chemical species will also be reversed, leading to an ―inverse‖ current peak.

Principles and concepts illustrated by this technique are: 1) quantitation of concentrations; 2) diffusion effects; 3) irreversibility; 4) study of reaction intermediates; 5) identification of electron transfer steps; and 6) relationships to the Nernst equation.[30] [31] [32] [33]

Voltammetric analysis are an important tool to help develop an amperometric biosensor.

1.1.1.2 Other transducer system

1.1.1.2.1 Piezo-electric biosensor

Piezo-electric biosensors are principally based on the measurement of change in resonant frequency of a piezo-electric crystal as a result of mass changes on its surface . These are caused by the interaction of tested species or analytes with a biospecific agent immobilized on the crystal surface

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and its resonant frequency changes as molecules adsorb or desorb from the surface of the crystal, obeying the relationships present in E 1.3:

E 1.3) A m Kf f 2  

Where Δf is the change in resonant frequency (Hz), Δm is the change in mass of adsorbed material (g), K is a constant for the particular crystal dependent on such factors as its density and cut, and A is the adsorbing surface area (cm2).

The frequency of vibration of the oscillating crystal normally decreases when the analyte binds to the receptor coating the surface. Such sensors generally operate by the propagation of acoustoelectric waves, either along the surface of the crystal or through a combination of bulk and surface, and are commonly referred to as surface acoustic (SAW) wave devices [34].

Most common piezoelectric biosensors are immunological biosensor including microgravimetric immunoassays, microbial assays, DNA hybridization, enzyme based detections and gas phase biosensors [35].

1.1.1.2.2 Optical biosensors

In the most commonly used form of an optical biosensor, the transduction process induces a change in the phase, amplitude, polarization, or frequency of the input light in response to the physical or chemical change produced by the biorecognition process Some of the advantages offered by an optical biosensor are selectivity and specificity, remote sensing, isolation from electromagnetic interference, fast, real-time measurements, multiple channels/multi parameters detection, compact design, minimally invasive for in vivo measurements, choice of optical components for biocompatibility, detailed chemical information on analytes, The main components of an optical biosensor are: light source, optical transmission medium (fiber, waveguide, etc.), immobilized biological recognition element (enzymes, antibodies or microbes, cells, protein, ect), optical detection system. The optical biosensors are classified in based at transduction methods (absorption, fluorescence, luminescence), geometry (bioptrode, Surface Plasmon Resonance, Fiber Grating Based Sensors) and Evanescent Wave Fiber Optic Biosensors.

The simplest optical biosensors use absorptions phenomenon to determine changes in the concentration of analytes. Chemiluminescence is same way similar to fluorescence. The difference is that chemiluminescence occurs by exciting molecules with a chemical reaction whereas fluorescence occurs by exciting molecules via light and then reemission of light at a longer wavelength [36].

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1.1.2 Immobilization methods in electrochemical biosensors

Biosensors often need that the biological material (enzymes, proteins, cells, tissues, nucleic acids) to perform its activity must be immobilized on the transducer of signal.

The choice of immobilization method depends on many factors, such as the nature of the biological element, the type of transducer, the physicochemical properties of the analyte and the operating conditions in which the biosensor is to function, and overriding all these considerations is necessary for the biological element to exhibit maximum activity in its immobilized microenvironment [37]. Generally there are 4 main immobilization methods: the adsorption, entrapment, covalent bonding and cross-linking immobilization.

The first one can roughly be divided into two others classes: physical adsorption and chemical adsorption. Physical adsorption is weak and occurs mainly via Van der Waals interaction. Chemical adsorption is stronger and involves the formation of covalent bonds. The physical adsorption is mostly used for enzyme immobilization in ZnO-based glucose biosensors.

The entrapment method refers to mixture of the biomaterial with monomer solution and then polymerised to a gel, trapping the biomaterial. However, this method can give rise to barriers to the diffusion of substrate, leading to reaction delay. Besides, loss of bioactivity may occur through pores in the gel. The gels commonly used include polyacrylamide, starch gels, nylon, silastic gels, conducting polymers, alginates etc.

In the covalent immobilization, the bond occurs between a functional group in the biomaterial to the support matrix. Some functional groups which are not essential for the catalytic activity of an enzyme can be covalently bonded to the support matrix. It requires mild conditions under which reactions are performed, such as low temperature, low ionic strength and pH in the physiological range.

In the cross linking method, usually, biomaterial is chemically bonded to solid supports or to another supporting material by a cross-linking agent. It is an useful method to stabilize adsorbed biomaterials. Glutaraldehyde is the mostly used bifunctional agent however the agents can also interfere with enzyme activity, especially at higher concentrations [19][38][39].

1.1.3 Biosensors to detect pesticides

In environmental pollution monitoring, it is becoming a general opinion that chemical analysis by itself does not provide sufficient information to assess the ecological risk of polluted waters and wastewaters. In the European Union, along with more stringent demands for water treatment (Council Directive 91/271/EEC), industrial and urban wastewater effluents shall reach certain limits

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of nontoxicity before the effluent can be discharged into the environment. Thus, much effort has been made during the last years to develop and use varius biosensors for toxicity evaluation of water samples [40].

The extensive use of pesticides for agricultural purposes is the cause of their widespread presence in natural waters. Concern about their toxicity and persistence in the environment has led the European Community to set limits on the concentration of pesticides in different environmental waters: The directive 98/83/EC on the quality of water for human consumption has set a limit of 0.1 μg/l for individual pesticides and of 0.5 μg/l for total pesticides.

To meet the demand for faster, sensitive and economic analytic methods to detect the environmental pollutants different biosensors have been developed, for example for the detection of organophosphorous and carbamate pesticides [41][42][43] by acetyl cholinesterase (AChE) and colin oxidase enzymes. Although sensitive, biosensors based on AchE inhibition are not selective (since the AchE is inhibited by neurotoxins, which include organophosphorous pesticides (OPs), carbamate pesticides, and many other compounds) and cannot, therefore, be used for quantitation of either an individual or a class of pesticides. Many others enzymes have been explored to developed biosensors to detect pesticides: the organophosphorous hydrolase (OPH) to detect organophosphate, [44], the tyrosinase and laccase to find phenols [45], diethyldithiocarbamates [46], and hydrazines [47]; Dithiocarbamate fungicides have been measured by their ability to inhibit the enzyme aldehyde dehydrogenase (AlDH) [48].

Photosynthesis inhibition is an interesting indicator that rapidly reflects the toxic effect of certain pollutants. Taking advantage of this feature, some biosensors based on Photosystem II (PSII) have been reported to be able to detect herbicides in the environment [49]. About 30 % of herbicides, including phenylurea, triazine, and phenolic herbicides, inhibit photosynthetic electron flow by blocking the PSII quinone-binding site and thus modify chlorophyll fluorescence [50]. Different amperometric biosensor are developed to detect different kind of herbicides too [51][10]. Heavy metals were also able to inhibit the activity of the PSII biosensor, but their effect is usually found at much higher concentrations than those typical for herbicides.

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1.2 Photosynthesis

1.2.1 Higher plants and green algae photosynthesis

In higher plants (and in the green algae) photosynthetic apparatus is represented by chloroplasts, special cell organellae which contain membrane-made closed structures (thylakoids) tightly leant one on the other to form piles, named grana (Figure 1.3).

Figure 1.3 A chloroplast. Thylakoids are packed in grana (is a stack of thylakoids folded on top of one another) or not. The stroma is the fluid space within the chloroplast. The lumen is the fluid filled space within a thylakoid.

Within thylakoid membranes the chlorophyll is associated in complexes containing up to 250 molecules, among which only very few are directly involved in the photochemical reactions producing ATP, most of them instead serving only as light-harvesting antennae. The function of these latter is just to collect and funnel light to the already mentioned [52][53] [54] [55] [56] . Such functional aggregates of a specific photosynthetic protein - as Photosystem II (PSII) for green plants - together with light harvesting complexes containing many chlorophyll and carotenoids with similar organization in both higher plants and photosynthetic bacteria [57].

The Photosynthetic apparatus is arranged in two photosystems, named Photosystem I (PSI) and II, (PSII, Figure 1.4) present in the thylakoids. They are an ensemble of several proteins and pigment-protein complexes working in concert building up functional units of metabolic importance. The final results of these interactions are: (1) the generation of a transmembrane proton gradient necessary to the ATP synthesis; (2) the production of the reducing power necessary to the NADPH biosynthesis [52] [54].

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Figure 1.4 The Photosynthetic apparatus is arranged in two photosystems, named Photosystem I (PSI) and II, (PSII) present in the thylokoids

The radiation absorbed by chlorophylls belonging to the light-harvesting complex of PSII is transferred to the pigment P680, made of two chlorophyll molecules arranged in such a way to form an excitonic dimer (the so-called ―special pair‖) and located at the interface between two similar subunits.

Within PSII light is absorbed by chlorophyll-protein harvesting complexes, then it is funnelled to the photochemically active reaction centre, made of D1, D2 and Oxygen Evolving Complex (OEC) subunits (Figure 1.5).

Figure 1.5 Representative PSII scheme, it is possible to see the principal protein D1 and D2, Oxygen Evolving Complex (OEC), Qa and Qb binding site

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Here photoexcitation converts a special dimer of Chl a molecules (P680, primary electron donor of PSII) , into its oxidized formP680+, thus triggering a single electron transfer first to a pheophytin cofactor, then to a plastoquinone molecule, Qa (located in D2) and finally to a second plastoquinone, Qbwithin D1 (Figure 1.5). Because of the homology in the structure and functions (e.g. for the sensitivity of both Reaction Center and PSII to triazine herbicides), the whole PSII is considered the ―eukaryotic analogous‖ of the bacterial photosynthetic Reaction Center (RC) [52], (Figure 1.6) whose properties will be discussed later.

a b

Figure 1.6 Homology in the structure between PSII and bacterial photosynthetic Reaction Center (RC)

The photooxided P680 dimer is then reduced by the electrons recovered from the oxidation of water to molecular oxygen operated by a Mn-containing complex bound to the reaction centre (Figure 1.5). Then the global reaction catalyzed by PSII is the light induced electron transfer from water - with the concomitant release of O2 to the second plastoquinone, together with the production of a transmembrane proton gradient.

The structured model [58] and the analysis of effects on the redox properties of cytochrome (Cyt) b559 induced by PSII herbicides [59] a third plastoquinone Qc was recently found, although its function is not fully clarified it could regular the redox potential in Cyt b559, which is characterized by an unusually high redox potential providing a mechanism of protection of PS II from photoinhibition. A representation of Qc and Qc binding sit is shown in Figure 1.7.

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a b

Figure 1.7 Schematic representation of the hypothetical collocation of Qc (a) and Qc binding site (b).

Subsequently electrons coming from PSII arrive to the PSI through the transmembrane cytochrome

bf complex (cytochrome complex in scheme), that moves protons inside the thylakoid space (the

space internal to thylakoid membranes of the grana) when electrons are transferred from PQH2 to plastocyanine (Pc) (a water soluble protein). At this point the thermodynamic driving force of the electron transfer is definitely exhausted: it is necessary a second photosystem, PSI, to promote a light-induced electron transfer from Pc through the pigment P700 (at the interface between identical subunits of the PSI) and then, through chlorophyll molecules, to ferredoxine (Fd) (a strong reducing agent).

Hence the ferredoxine-NADP+ reductase, a flavoprotein located on the stromal side of the membrane, will catalyze the NADPH synthesis. By this way the connection between PSII and PSI allows the electron transfer from H2O to NADPH to proceed and, moreover, it leads to the formation of a proton gradient necessary for the ATP synthesis.

The global reaction of the passage through PSII, Cyt bf and PSI follows (R 1.5):

R 1.5) 2NADP2H2OLIGHT  2NADPH22H O2

Alternatively, electrons can be transferred from Fd to PSI through the cytochrome bf complex, instead of going to NADP+: this cyclic photophosphorylation leads to production of the proton gradient without NADPH synthesis (as in bacteria with NADH). This biochemical shunt is active when no NADP+ is available: following this pathway, PSII is not involved in the photosynthetic process and therefore no oxygen production is observed. Subsequently ATP and NADPH so formed

19

other organic compounds, through the ―dark phase‖, the so-called Calvin-Benson cycle, whose global reaction is [52] (R 1.6):

R 1.6) CO22H2O3ATP2NADPH





It is possible summarize the reaction R5 and R6 in with the basic follow equation where CHOH is carbohydrate unit (Reaction R 1.7):

R 1.7) CO2H2OLIGHT 





1.2.2 Purple Bacteria

Purple bacteria are unicellular prokariotes: they are phototrophic since they are able to use solar energy to oxidize reduced substrates, being carbon dioxide the final acceptor of the electrons, they are divided in Thiorhodaceae or Chromatiaceae (purple sulphur bacteria); or Athiorhodaceae or Rhodospirillaceae (purple non-sulphur bacteria) [60] depending on whether they use sulphur compounds as reducing agents for photosynthetic carbon dioxide fixation (water for high plant and green algae). For our purposes, we work with Rhodobacter sphaeroides part of Rhodospirillaceae, many of them are facultative anaerobes, but lose most of their photosynthetic pigmented proteins when they are grown aerobically (which may be in the dark or not). In the presence of oxygen, the cells of the Rhodospirillaceae carry out respiration, behaving like eukaryote‘s mitochondria.

They constitute the largest group of photosynthetic bacteria and account for 8 genera and 30 species identified up to now. To this group belongs, rod-like bacterium provided with flagella.

Purple bacteria carry out a photosynthetic process called anoxygenic, (no produce oxygen). They are able to utilize either the products of the anaerobic metabolism of the sea depths (hydrogen sulfide, hydrogen gas, carbon dioxide) or the product of the anaerobic fermentations and of the partial oxidations taking place in the intermediate layers of the stagnant waters (lakes, pounds, lagoons). For this reason they are able to grow even in the dark, aerobically oxidizing the reduced carbon compounds, even though their optimal oxygen concentration is low. Their native environment is limited to the water layers closer to the surface, at the border between oxygen-containing and oxygen-free waters, where the oxygen concentration is low.

As already specified, the anoxygenic photosynthesis of purple bacteria is different from the oxygenic one of green plants. In prokaryotes there are no chloroplasts and the photosynthetic

20

pigments are localized just on the plasmic membrane organized in structures deriving from its extensive folding.

R.sphaeroides photosynthetic apparatus is formed by two main integral-membrane enzymes, the

Light Harvesting Complex (LH, type I and II), harvesting the electromagnetic radiation, and only one RC, that utilizes the light absorbed (only one compared to the two of the green plants, one for each photosystem) (Figure 1.8).

Figure 1.8 LH type I and II harvest the electromagnetic radiation to RC that utilizes the light absorbed to photosynthesis

A single RC molecule is made of up to 25÷30 bacteriochlorophyll molecules, although only very few of them are directly involved in the photochemical reactions leading to the ATP synthesis. The RC is composed of three subunits termed H (heavy), M (medium) (corresponding to D2 protein of PSII in higher plant and green algae) , and L (light) (corresponding to D1 protein o of PSII in higher plant and green algae) on the basis of their migration properties in sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis (PAGE) (Figure 1.6b and 1.9a).

The three subunits are present in a 1:1:1 stoichiometry and contain a Bchl a dimer, two monomeric molecules of Bchl a, two monomeric molecules of bacteriopheophytin a, two ubiquinones Qa and Qb, and a nonheme iron [61] (Figure 1.9b). The binding sites for each quinones were present in H and L protein respectively, as the high plant homologues protein D2, D1(Figure 1.6).

21

Figure 1.9 Structure, mechanism and environment of the R. sphaeroides RC. (a) The ten cofactors (spheres) are held in place by a scaffold consisting of L and M polypeptides (ribbons). The H-polypeptide has a single TM helix and a cytoplasmic domain. (b) The BChl, BPhe and ubiquinone cofactors.

Purple bacteria do not produce oxygen and do not use water us external electron donor as explained in the reaction R 1.8 where H2A and A are the external not specific electron donor in according to the available environmental sources. and to the bacterial type. They can be H2S and S, or H2 and CHOH it is the carbonit unit) [62]:

R 1.8) CO22H2ALIGHT 





The Calvin-Benson cycle, utilizing as reducing agents the NADPH or the NADH coenzyme, besides ATP for energetic needs. The NAD+ reduction to NADH is performed by the NADH dehydrogenase, operating in an opposite direction in relation to that of the cell respiration: the electrons are extracted from the reduced substrates (H2A) and that represents the conclusions of the light-dependent part of the bacterial photosynthesis, whose complete reaction – in analogy with the equation R 1.9:

R 1.9)2NAD 2H2ALIGHT  2NADH22A 2H

The light, harvested by LHCI and LHCII, is funnelled to the RC primary photoinduced electron donor (a bacteriochlorophyll dimer), the RC being another of the three complexes. An electron is therefore raised to an energy level suitable to initiate the electron transfer that, through a series of intermediate acceptors, reaches the final acceptor, usually an ubiquinone molecule. Once fully reduced - i.e.: after taking up two electrons as well as two protons from the cell cytoplasm - the

22

ubiquinol can leave the RC exchanging with a pool of ubiquinone molecules present in the core of the bacterial membrane (Figure 1.10). The reduction of a single ubiquinone molecule requires, therefore, the absorption of two photons by the RC. This means that at the end of the first turn of the photocycle (leading to the formation of a semiquinone radical), the photooxidized bacteriochlorophyll dimer needs to be reduced.. To close the cycle the cytochrome (cyt) bc1 (the last of the three complexes of the photosystem) oxidizes the ubiquinol just arrived from the membrane quinone pool, with the concomitant release of two protons in the periplasm. As a consequence, results of the absorption of two photons is the traslocation of two protons from the cytoplasmic to the periplasmic side of the bacterial cell. Within this perspective, RC can be assimilated to a light driven pump producing the proton gradient required for the right function of both the ATP synthase - the third photosynthetic membrane complex – and the NAD+- reductase. Purple bacteria are also able to vary their proportions of light-harvesting to RC complexes in response to changes in the level of light [62] - as it happens for the extent of membrane invagination. In the high-light conditions, moreover, higher amounts of quinone (electron acceptor), of cyt bc1 complex (electron transporter), and of ATPase particles have been found, always in relation to RC proteins: in these conditions the arrangement of the antenna complexes in the membranes is modified, smaller units would be more efficient and the RC turns over more often, allowing faster electron transfer within the photosystem. Size and efficiency of the photosystem are regulated by light intensity, the synthesis of the whole photosynthetic apparatus has been found to be controlled by oxygen at a transcriptional level [62].

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Figure 1.10: A schematic representation of cyclic-anoxogenic photosynthesis in R.sphaeroides

1.3 Photosynthesis biosensor

In recent years concern for drinking water contamination from industrial and agricultural activities has been rising. Consequently, the need for efficient environmental monitoring strategies became urgent. Monitoring devices have to be rapid, easy to operate, and offer low-cost screening procedures possibly applicable on field.

Even low levels of chemicals may cause strong adverse effects on humans, animals, plants and ecosystems.

Herbicides represent a specific class of pesticides employed in the chemical weeding of various crops. The most commonly used in agriculture (about 30 % of the total) are urea derivatives, triazines, diazines and phenols, acting as inhibitors of photosynthesis [63]. In particular, triazines (atrazine, simazine, terbuthylazine etc.) and ureas (linuron, diuron, etc.) from agricultural runoff can contaminate soils, surface and ground waters with severe toxic effects on humans. Moreover, their widespread use may lead to the natural selection of resistant weed species, in turn requiring higher amounts of herbicides in agriculture. Although the use of atrazine and simazine was banned by law in European Union (EU) in 2004 (after a 12-year review period) [63][64]. their persistence in soil and water is many years- long due to the combination of a very low water solubility and difficulties in being metabolized by microorganisms [65].

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Moreover, up to date the European Commission has published a priority list of substances, which is setting up a framework for an action in the field of water policy, in order to preserve, protect and improve the quality of environment (2455/2001/EC Water Framework Directive, WFD). According to WFD, an efficient monitoring of priority pollutants is required, together with assessment of biological/ecological quality elements and measurement of physic-chemical parameters. Since then developing biosensor devices for rapid and accurate prescreening and determination of pesticides in water samples became a crucial objective.

Biosensors are designed to match many scientific, medical, industrial and environmental applications due to the advantage of being generally sensitive, rapid, economical and easy-to-handle for on-field measurements (see above) [66], from 1980s onwards many biosensors focused on photosynthetic processes. Representing the heart of photosynthesis, the Photosystem II (PSII) multi-subunit pigment protein complex is embedded within thylakoidal membranes of plants, algae and cyanobacteria. PSII. Light triggers an intraprotein electron transfer chain proceeding also beyond PSII, up to other photosynthetic proteins such as cytochrome bf, plastocyanine and Photosystem I (PSI), until the final double electron reduction of NADP+ to NADPH.

Among several advantages, the usefulness of using biosensors based on photosynthetic particles containing PSII is due to the fact that the D1 protein specifically recognizes and binds herbicide molecules within its so called Qb binding site, where they act as inhibitors of the photosynthetic activity in competition with a native plastoquinone cofactor. Depending on their concentration, herbicides partially or fully block the electron transfer between Qa and Qb quinones, leading to interruption of water photolysis and oxygen production (Hill reaction) [67] the inhibitor‘s effect depends on the structural conformation of the Qb site, in turn depending on the arrangement of the D1/D2 subunits. This binding property has paved the way for the use of PSII as an analytical tool for the design and detection of herbicides [68] [69]. Biosensors based on photosynthetic particles containing eukaryotic PSII complexes, as well as their prokaryotic precursors bacterial Reaction Centre (RC) proteins, from whole cells to photosynthetic organelles and isolated proteins of various organisms (algae, cyanobacteria, purple bacteria and green plants such as spinach) have been reported to be able to detect triazine, diazine, urea and phenol pesticides in water samples, both by optical [70][71][72][73], and electrochemical transduction systems [10][74][75][76][77]. Although a variety of whole-cell-based bacterial sensors have been applied in environmental assays for pollutant monitoring, the molecular details of RCs bacterial compared to PSII D1/D2 subunits (analogous in eukaryotes, with very high sequence homology) make RCs biosensors generally less sensitive to herbicides in comparison to algal and cyanobacterial basel devices [72][73]. It has been

25

shown that photosynthetic organelles such as thylakoidal membranes (TMs) from green plants [10][67][70], have the feature of selectively recognizing herbicides and remaining active once extracted from their native organism. Nevertheless, features such as stability and analytical sensitivity toward pesticides of TMs-based biosensors, very similar to those of whole algal cells biodevices, together with time-wasting extractions of organelles, have not encouraged their diffusion as potential commercial instruments [10] [78].

1.4 Aim and develop of PhD project

An innovative amperometric biosensor equipped with flow injection system was designed and realised for the detection of pesticides in water samples. It relies on the well-known ability, exhibited by the photosynthetic PSII protein complexes of selectively binding some classes of pesticide compounds such as triazine and ureic herbicides, within the so-called quinone Qb site of PSII. Therefore there is a competition between the native PSII cofactor of the Qb site, a quinone, and the added herbicide molecules (see above)

The aim of this PhD project, within the European Project ―Bio-sensor for Effective Environmental Protection and Commercialization - Enhanced‖ (BEEP-C-EN), is to enhance the sensitivity and the stability of the photosynthetic-based biosensors to identify different herbicides classes and subclasses in the environment and agrofood. In this project two different photosynthetic microorganisms will be used: a green algae (Chlamydomonas reinhardtii) and a purple bacterium (Rhodobacter sphaeroides). These microorganisms were chosen for the sensitivity towards herbicies (C. reinhardtii) [10] [72] [79] and stability (R. sphaeroides) [73] [80] of their reaction centres.

Two parallel approaches were undertaken: 1)developing a new amperometric biosensor using whole cell algae as biosensing element, and 2) a genetic engineering to realize a Reaction Center chimera protein as a new biomediator potentially joining the sensitivity feature of the green alga plus the stability feature of the purple bacterium.

1.4.1 Development of biosensor whole cell based C. reinhardtii

Although thylakoid membranes extracted from spinach leaves and suitably immobilized over electrodes have been revealed to be quite stable (half-life of 9 h), reusable and very sensitive to herbicides (limit of detection of  1*10-8 M with most common triazine and urea compounds) we showin this PhD thesis that similar and even better results may be reached by means of algal cells

26

immobilized over cheap screen-printed electrodes (SPEs). Compared to thylakoid membranes, algal cells present the strong advantage of requiring no time-consuming extraction from photosynthetic organisms except a simple cell concentration by centrifuge nor the use of electrochemical mediators such as Tetramethyl-p-benzoquinone (duroquinone) or 2,6-dichlorophenol-indophenol (dichlorindolphenol) for amperometric measurements[10].

We selected as most versatile photosynthetic biomediator for our amperometric measurements were

Chlamydomonas reinhardtii algae cells.

On the basis of recent literature [81] [82] [83] [84] [85], giving priority to easy and biocompatible (i.e. compatible to algal biomediator) immobilization procedures, we decided to adopt and compare only two simple techniques among those above mentioned: (a) coating of algae cells with a Nafion® (Figure 1.11a) membrane and (b) physical gelation of alginate (Figure 1.11b), entrapping algae cells.

Nafion® , that is a sulfonated tetrafluoroethylene based fluoropolymer-copolymer discovered in the late 1960s by Walther Grot of DuPon [86]. In biosensor field, this polymer was often used to immobilize enzymes, nucleic-acids, and other macromolecules [87] [88], but it was never used to immobilize photosynthetic whole cell algae.

Another of the most promising encapsulation material, already used for algae immobilisation, is the alginate hydrogel [78]. Alginates, extracted from brown seaweeds, are linear polysaccharide copolymers of (1-4)-covalently linked β-D-mannuronate and α-L-guluronate

monomer residues arranged in different sequences (Figure1.11b). Due to their abundance, lack of toxicity and compatibility with biological systems, alginates are widely used for immobilization procedures [89] [90] [91]. Gelation of alginate is achieved by an ion exchange between sodium from the guloronic acid salts and divalent cations such as Ca2+. Unique properties of alginates are: (a) the possibility of exploiting a room temperature gelation/encapsulation process; (b) a relatively inert aqueous environment within gels; (c) high gel porosity, allowing high diffusion rates for macromolecules. These properties have enabled them to be used as matrices for the entrapment of several proteins.

.

27

With these two different immobilization techniques, a biomediator to detect pesticides was developed optimizing the experimental condition to enhance its stability and sensitivity.

1.4.2 Development of Chimera RC Protein

The first experimental approach to create a Chimeric RC was to check the feasibility of the “L-D1

loop” by bioinformatic study with the support Prof. Fabio Polticelli of the Computational

Biochemistry Lab, University of Roma3. After the bionformatic study , I produced the Chimera in the laboratory of Biochemistry School, Medical Science of University of Bristol under the supervision of Mike R. Jones. The Chimera was created, and further will be tested as biomediator to develop a new biosensor for detection of low concentration of pesticides able to control the limits imposed in the European Country.

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2.Experimental Section

2.1 Development of biosensor based on whole cells of Chlamydomonas reinhardtii

2.1.1 Chlamydomonas reinhardtii growth

C. reinhardtii is a soil organism, it can be grown in laboratory either in liquid culture or on agar in

simple mineral salts, and many of the recipes proposed for algae are in general sufficent for its culture.

C. reinhardtii cultures, all strains, IL (intron less) [92] F274Y(Phenylalanine in position 274 was substituted with a Tyrosine), were grown in liquid Tris–Acetate-Phosphate (TAP) medium pH 7.0÷7.2 in 250 ml flasks, at 50 µmol/m2s light intensity, 25°C temperature and 150 rpm stirring. [78]. When required, this medium was solidified with 1.5 % w/v agar to prepare Petri dishes to have ―solid‖ culture (Figure 2.1b), able to store the algae for about a month. After this period usually the dishes need to be plated again. The plate is useful to produce the liquid culture: a small algae quantity is taken from the dish with a sterile handle and inserted in a Erlenmeyer flask containing the medium (Figure 2.1a) and grown in agitation as above described.

a b

TAP medium [93] is well buffered and relatively low in phosphate, making it the best medium for 32P labeling and for studies requiring optical clarity of agar, but it is the most expensive of the usual media to prepare. TAP medium contains 17.4 mM acetate as originally formulated. A Figure 2.1 Example of the liquid (a) and solid algae culture (b)

29

minimal medium can be made by omitting acetic acid and titrating with HCl to pH 7.0. In addiction of TAP the ―trace elements‖ (as EDTA, citrate, iron salts etc.) are added, asthe most used trace elements developed by Hutner et al. (1950) [94]. It is possible to see the production in Protocol Section, Protocol 1.

2.1.2 Chlamydomonas reinhardtii characterization

Before immobilization, algae cells withdrawn from culture flasks within their early, mid-exponential growth phase (0.4÷0.6 absorbance at 750 nm) were always characterized in terms of chlorophyll content, cell density and fluorescence yield to ensure reproducible procedures on viable cells.

2.1.2.1 Chlorophyll content.

In according with Porra et al. (1989) [95] the total chlorophyll concentration was calculated: 1 ml of culture was centrifuged at 2000 g for 2 min (Heraeus Biofuge Pico tabletop centrifuge, Kendro, Newport Pagnell, UK). 800 μl of supernatant were discarded and substituted by 800 μl of pure acetone (Carlo Erba, HPLC grade, 99.9 % minimum purity), then the resulting mixture was vortexed for 2 min and centrifuged for 4 min at 13800 g to eliminate the cell starch present as pellet. The chlorophyll content in the supernatant, referred to 1 ml of original culture (μg/ml), was determined spectrophotometrically at 652 nm against 80 % V/V acetone/water, according to the formula (Abs652nm×1000)/34.5 [96]. Usually values as 10÷15 μg/ml were obtained.

2.1.2.2 Cell density.

When cultures showed Abs750= 0.4÷0.6 [97], the cells were daily counted with a Bio-Rad TC-10 automated cell counter (Hemel Hempstead, UK) providing an average cell density of (8±2)*105 cells/ml.

2.1 2.3 Fluorescence yield.

The cell viability was evaluated also by measuring the chlorophyll fluorescence induction curves with a Plant Efficiency Analyzer instrument (PEA, Hansatech Instr. Ltd, Kings Lynn, Norfolk, UK) on 5 ml algal culture samples suitably diluted (OD750 1) after 10 min of dark-adaptation. The excitation light, a 5 s saturating pulse (600 W/ m2), was provided by an array of six red light emitting diodes (LEDs with maximum emission at 650 nm) focused on the sample surface. As

30

indicator of the cell photosynthetic performance the maximum quantum yield of PSII photochemical reaction, Fv/Fm= (Fm-F0)/Fm, was used, where F0 and Fm indicate the minimum (50

µs after the onset of the excitation light) and Fm fluorescence intensities, respectively. The variable fluorescence, Fv, is calculated as Fm-F0 and represents the fraction of photosynthetically active PSII

[98].

2.1.2 Algae cells Nafion® immobilization on screen-printed electrodes

C. reinhardtii cells were concentrated by 2 minutes centrifugation at 15400 x g and the pellet was

washed for 2 times with immobilization buffers (2 or 3, respectively composed by Tricine 20mM-NaOH pH 7.2, Sucrose 70mM, NaCl 50mM, MgCl2 5mM, Ac-K 2mM, and Tricine 20mM-NaOH pH 7.2, Sucrose 70mM, NaCl 10mM, MgCl2 5mM, Ac-K 2mM ) by 2 minutes centrifugation at 15400 x g. The suspension of cells obtained was spotted on the SPE (WE) and left at 4°C in the dry box for few minutes until it were dried. After that, 1% Nafion® solution (prepared in immobilization buffer) was spotted onto the biologic material, and dried following the same procedure mentioned before. On each SPE, 224500 cells were immobilized.

2.1.3 Algae cells Alginate immobilization on screen-printed electrodes

An aliquot of C. reinhardtii culture in its exponential phase growth (usually within 10÷18 ml and characterized as above reported), corresponding to 1.2*107 cells was collected and centrifuged at 2000 x g for 5 min at 15°C with a refrigerated ALC PK121R centrifuge (UK). : the withdrawal operation has been standardized also on the basis of culture Abs750 values, with similar results. Withdrawals from cell cultures standardized on the basis of 750 nm absorbance provided the same results [6]

After washing the supernatant was discarded, while the pellet was resuspended with Tricine-NaOH 50 mM pH 7.2 to a final volume of 50 µl, then mixed with 100 µl of 2 % w/V sodium alginate solution in the same buffer, obtaining 150 µl of viscous algae/alginate suspension. 5 µl of the suspension (containing 4*105 cells, see Results) were dropped over a Multi-Walled Carbon Nanotube (CNT) working electrode surface (diameter 4.0 mm) of a commercial Screen-Printed Electrode (SPE) (DRP-110CNT, DropSens, Oviedo, Spain, counter and reference electrodes were made of CNT and Ag/AgCl respectively). Then the CNT-SPE (Figure 2.3c) with spotted material (biomediator) was immersed for 20 minutes in Tricine-NaOH 50 mM, CaCl2 200 mM (Gelation buffer) prior to use, in order to obtain a physical gelation of the alginate polysaccharide and

31

consequent entrapment of the biomediator on the working electrode. A single preparation of 150 µl suspension provided over 20 SPEs bearing the immobilised biomediator (hereafter referred also as screen-printed biosensors, SPE-Chlamy) took  40 min including gelation time.

2.1.4 Biosensor instrument setup

The whole measuring setup was made of 3 components: (1) An electro-mechanical device equipped with suitable software (AMPBIO-SPE instrument developed by Biosensor s.r.l., Rome, Italy) coupled to a peristaltic pump [99] and a potentiostat (PG581 model, Uniscan Instruments Ltd, Buxton, UK) (Figure 2.2). (2) A disposable screen-printed cell with classical three electrodes design: screen-printed CNTs as working electrode, Ag/AgCl as reference electrode and an auxiliary CNTs electrode. (3) (Figure 1.2 and 2.3) An immobilized photosynthetic material (unicellular algae) responsible of the target analyte recognition. The AMPBIO-SPE is a portable instrument specifically designed for electrochemical measurements on biological samples immobilized on disposable SPEs suitably connected to the potentiostat.

Voltammetric and amperometric measurements on SPE whit the immobilized algae (SPEs-Chlamy) were carried out using the above mentioned apparatus. For this purpose SPEs-Chlamy were easily inserted upside down through a slit into a leakproof flow chamber made of Delrin® (polyoxyethylene), 30 mm wide in diameter and 25mm high (Figure 2.2 bc). Two red LEDs (λ= 650nm, 325 μmol/m2s1 total light intensity) positioned below the transparent bottom surface of the cell stimulate the photosynthetic activity of the biomediator during electrochemical measurements. Amperometric tests were carried out in dynamic mode with a suitable buffer solution flowing at rate of 0.1 ml/min through the measurement chamber by means of a peristaltic pump equipped with silicone (Masterflex) tubes, (lying downstream from the chamber and working in aspiration mode). The output tube from the measurement buffer solution container and upstream from the chamber instead was made of chemically inert polytetrafluoroethylene (Teflon). The whole apparatus was equipped also with electronic control board for data read-out, processing and storage. Programmable parameters were LED intensity and time of illumination (light/dark sequence), flow rate, and other than specific electrochemical parameters. Keyboard and display can be used to set the parameters, while a USB connection allows data transfer to PC.

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2.1.5 Amperometric Measure

2.1.5.1 Electrode Choosing

Different screen printed electrodes (SPEs) with different working electrode (WE; graphite, graphite nanotube, gold Figure 2.4) were used and tested.

a) b)

c)

Figure 2.2 a) Biosensor Instrument is shown with all its key stakeholders: of a potentiostat PG581 (Uniscan Instrument Ltd) (a) the a peristaltic pump into a 70µl chamber containing on the button two red leds (650nm, 1500mcd) (b)(c). In c, a particular of the cell and the right positioning of the SPE-Chlamy.

a) b) c)

d)

Figure 2.3 - Example of Screen Printed Electrode: (a)carbon SPE, (b) carbon nano-tube one (CNT, the difference between a and b working surface composition is illustrated in Figure d by SEM analysis,

33

These WE were purchased from by our partner of the European project Dropsens (http://www.dropsens.com, Oviedo Asturias, Spain) .With the cyclic voltammetry (CV) the best electrode usable at our work potential (-0.7V) [81][82] (Figure 2.4) was chosen.

2.1.5.2 Setting of Electrochemical Measures (Nafion® immobilization)

A set of buffers was tested as electrochemical buffer to find out which gave the best and stable signal over time. The buffers used were: buffer 1 composed of Tricine-NaOH pH 7.2 20mM, Sucrose 70mM, NaCl 50 mM, MgCl2 5mM ; buffer 2 composed of Tricine 20mM, Sucrose 70mM, NaCl 50 mM, MgCl2 5mM, potassium acetate (Ac-K) 2mM; buffer 3 composed of Tricine 20mM, Sucrose 70mM, NaCl 10 mM, MgCl2 5mM, Ac-K 2mM; buffer 4 composed of Tricine 20mM, Sucrose 70mM, NaCl 10 mM, MgCl2 5mM; and buffer 5 composed of Tricine 20mM, Sucrose 70mM, MgCl2 22mM. Some of these buffers were used as immobilization buffers, or measure buffer or for both purposes. In this initial experiment only the IL strain was tested in different conditions (using different buffers) Herbicides inhibition of electrons transport was verified.

For Chronoamperometry analysis (CA) the potential was set at -0.7 V vs. Ag/AgCl according to Shitanda [82], at this potential the oxygen is reduced.

From two references 1 min light followed by 4 min darkand 15 s light/45 s dark sequences are drawn as staring conditions. Subsequently, in order to allow a longer dark relaxation time, a sequence of 1 min light/5 min dark has been adopted. After that a careful observation of time evolution for amperometric signals under illumination clearly showed that a light interval of only 30 s was enough to reach the maximum height of the peak, the following 30 s being unnecessary. Figure 2.4- Cyclic Voltammetry comparison of three different SPEs, shows the carbon-nanotubes is the most suitable on the biomediator work potential (-0.7V)